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model for multiple sclerosis
Nathalia Teixeira, Gisele Picolo, Aline Giardini, Fawzi Boumezbeur, Géraldine
Pottier, Bertrand Kuhnast, Denis Servent, Evelyne Benoit
To cite this version:
Nathalia Teixeira, Gisele Picolo, Aline Giardini, Fawzi Boumezbeur, Géraldine Pottier, et al..
Al-terations of peripheral nerve excitability in an experimental autoimmune encephalomyelitis mouse
model for multiple sclerosis. Journal of Neuroinflammation, BioMed Central, 2020, 17 (266), pp.1-14.
�10.1186/s12974-020-01936-9�. �hal-02933447�
R E S E A R C H
Open Access
Alterations of peripheral nerve excitability
in an experimental autoimmune
encephalomyelitis mouse model for
multiple sclerosis
Nathalia Bernardes Teixeira
1,2,3, Gisele Picolo
2, Aline Carolina Giardini
2, Fawzi Boumezbeur
3, Géraldine Pottier
4,
Bertrand Kuhnast
4, Denis Servent
1and Evelyne Benoit
1*Abstract
Background: Experimental autoimmune encephalomyelitis (EAE) is the most commonly used and clinically relevant murine model for human multiple sclerosis (MS), a demyelinating autoimmune disease characterized by mononuclear cell infiltration into the central nervous system (CNS). The aim of the present study was to appraise the alterations, poorly documented in the literature, which may occur at the peripheral nervous system (PNS) level.
Methods: To this purpose, a multiple evaluation of peripheral nerve excitability was undertaken, by means of a minimally invasive electrophysiological method, in EAE mice immunized with the myelin oligodendrocyte glycoprotein (MOG) 35-55 peptide, an experimental model for MS that reproduces, in animals, the anatomical and behavioral alterations observed in humans with MS, including CNS inflammation, demyelination of neurons, and motor abnormalities. Additionally, the myelin sheath thickness of mouse sciatic nerves was evaluated using transmission electronic microscopy.
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© The Author(s). 2020 Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visithttp://creativecommons.org/licenses/by/4.0/. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated in a credit line to the data.
* Correspondence:evelyne.benoit@cea.fr
1
Université Paris-Saclay, CEA, Département Médicaments et Technologies pour la Santé (DMTS), Service d’Ingénierie Moléculaire pour la Santé (SIMoS), ERL CNRS 9004, Gif-sur-Yvette, France
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Results: As expected, the mean clinical score of mice, daily determined to describe the symptoms associated to the EAE progression, increased within about 18 days after immunization for EAE mice while it remained null for all control animals. The multiple evaluation of peripheral nerve excitability, performed in vivo 2 and 4 weeks after immunization, reveals that the main modifications of EAE mice, compared to control animals, are a decrease of the maximal compound action potential (CAP) amplitude and of the stimulation intensity necessary to generate a CAP with a 50% maximum amplitude. In addition, and in contrast to control mice, at least 2 CAPs were recorded following a single stimulation in EAE animals, reflecting various populations of sensory and motor nerve fibers having different CAP conduction speeds, as expected if a demyelinating process occurred in the PNS of these animals. In contrast, single CAPs were always recorded from the sensory and motor nerve fibers of control mice having more homogeneous CAP conduction speeds. Finally, the myelin sheath thickness of sciatic nerves of EAE mice was decreased 4 weeks after immunization when compared to control animals.
Conclusions: In conclusion, the loss of immunological self-tolerance to MOG in EAE mice or in MS patients may not be only attributed to the restricted expression of this antigen in the immunologically privileged environment of the CNS but also of the PNS.
Keywords: Electrophysiology, Experimental autoimmune encephalomyelitis, Mouse, Multiple sclerosis, Myelin oligodendrocyte glycoprotein, Neuromuscular junction, Peripheral nervous system
Introduction
Multiple sclerosis (MS) is a chronic, inflammatory, and demyelinating disorder of the central nervous system (CNS), considered as an important and frequent neuro-logical impairment condition [1, 2]. It is a disorder of autoimmune origin, where the immune system recog-nizes parts of the CNS as antigens, specifically peptides that form the myelin sheath of axons of neurons [3–5] leading to a demyelination process which induces ser-ious physical, cognitive, emotional, and social problems [1, 2]. Although the evolution of the disease is highly variable, the disability faced by most people is irrevers-ible. Therefore, MS is considered incurable, and the dif-ferent therapeutic options focus on delaying disease progression and promoting the relief of symptoms so as to maintain the quality of life of patients [2]. In contrast to therapies focused on controlling or modulating the immune (innate and adaptive) responses to limit demye-lination and neuronal damage, novel drugs currently in clinical trial have been recently reported to promote re-pair and regeneration in the CNS [6].
The immunopathogenesis of MS is not completely under-stood, and it sets up as a picture that remains to be eluci-dated. However, important steps are clearly involved in this disease, primarily at the CNS. Both innate and adaptive im-mune system responses are dysregulated in MS [7]. It is a mainly T cell-mediated disease with a central role of myelin-reactive CD4+ T cells [8, 9]. Autoreactive T-helper type 1 (Th1) and 17 (Th17) cells are peripherally activated and sub-sequently migrate to the CNS, causing central inflammation with release of cytokines, microglial activation, axonal and myelin injury, followed by demyelination and atrophy of white matter tract across the brain and spinal cord [10–12],
which causes neurological and motor impairment. The atro-phy occurs in key regions. Posterior cingulate cortex, precu-neus, and thalamus are among the earliest regions to become atrophic [13]. In addition to the neuroinflammation, some other factors may underlie the neurodegeneration and brain atrophy, including mitochondrial failure, iron depos-ition, and retrograde neurodegeneration in the deep gray matter through white matter lesions [14–16].
B cells and antibodies have also a role in the pathology of MS. High levels of immunoglobulins in the cerebro-spinal fluid of patients were detected together with an increase in these levels during periods where the symp-toms were worse [17–19]. The production of myelin-specific antibodies (and the consequent rupture of mye-lin sheets) seems to be an important way in which B cells contribute to the disease [19]. Recently, a more central role of B cells in MS, which appears to be anti-body independent, has been described [20, 21]. Accord-ing to this recent scenario, B cells would activate or downregulate the proinflammatory responses of both myeloid and T cells, and recruit autoreactive T cells to the CNS. Thereafter, interactions among these cells would determine the development of MS episodes.
In contrast to the well-defined and critical central effect, the peripheral alterations, although frequently de-scribed in patients, have not been characterized as thor-oughly [22–25]. In particular, histological studies performed on rats with experimental autoimmune en-cephalomyelitis (EAE) showed that inflammation was present in both peripheral nervous system (PNS) and CNS [22]. In this model of EAE induced by passive transfer of a cytotoxic CD4+ T cell clone specific for the 72-89 peptide of guinea-pig myelin basic protein (MBP),
the spinal roots in the PNS as well as the spinal cord root entry and exit zones in the CNS were considered as the main sites of demyelination at periphery. More re-cently, electrophysiological analysis revealed a mem-brane hyperexcitability of sensory neurons of dorsal root ganglia isolated from MOG35-55-induced EAE mice, pro-viding evidence of peripheral sensitization in MOG-EAE murine model [23].
The frequency of peripheral demyelination in a whole MS population is unknown, although some case reports have been described to be associated with demyelinating neuropathy [26,27].
Few studies have investigated peripheral alterations in-duced by MS. Hence, peripheral sensory and motor ab-normalities were analyzed in 20 patients showing MS by evaluating conduction velocities and amplitudes of ulnar, sural, and tibial nerves. Electrophysiological abnormalities were found in 15 of 91 nerves examined (16.5%) but neurological disability was not associated with the pres-ence of electrophysiological abnormalities [28]. In addition, nerve conduction abnormalities suggestive of de-myelination were demonstrated in only 5% of 60 patients with relapsing-remitting MS [29]. This later study, in con-trast to a number of case reports describing patients with MS who develop demyelinating neuropathy, strongly sug-gests that central and peripheral demyelination coexist only in a special subgroup of patients with MS.
The aim of the present work was to study the periph-eral impairment induced by MS. To this purpose, peripheral sensory and motor nerve excitability was eval-uated in a MOG35-55-induced EAE mouse, an experi-mental model for MS that reproduces, in animals, the anatomical and behavioral alterations observed in humans with MS, including CNS inflammation, demye-lination of neurons, and motor abnormalities [3,30–32].
Materials and methods
Animals
Animal experiments are reported in line with the AR-RIVE (Animal Research: Reporting of In Vivo Experi-ments) guidelines developed in consultation with the scientific community as part of an NC3Rs initiative to improve standards of reporting the results of animal ex-periments, maximizing information published, and min-imizing unnecessary studies [33, 34]. Experiments were carried out on 8-week-old female C57BL/6 mice (Mus musculus, weighting 18–20 g) purchased from Janvier Elevage (Le Genest-Saint-Isle, France). We used exclu-sively female mice taking into account that first, MS is an autoimmune disease where the incidence is higher in women than in men [35–37] and second, most of the previous studies were performed on female animals. The animals were acclimatized at the CEA animal facility for at least 48 h before experiments, and were treated in
strict adherence with the European Community lines for laboratory animal handling and to the guide-lines established by the French Council on animal care “Guide for the Care and Use of Laboratory Animals” (EEC86/609 Council Directive—Decree 2001-131). In particular, the mice were housed in a room with con-trolled temperature and a 12-h light/12-h darkness cycle, in standard laboratory cages with environmental enrich-ment (bedding and cardboard tubes), and were allowed to free access to water and food.
All animal experimental procedures were approved by the Animal Ethics Committee of the CEA, by the French General Directorate for Research and Innovation (pro-ject APAFIS#5973-2016070515456532v6 authorized to FB and project APAFIS#2671-2015110915123958v4 au-thorized to EB) and by the Butantan Institute (CEUAIB protocol number 7334170718 authorized to GP).
EAE mouse model
EAE was induced as previously described [38, 39]. Briefly, each EAE mouse was immunized with 200μg of synthetic myelin oligodendrocyte glycoprotein (MOG) 35-55 peptide (MEVGWYRSPFSRVVHLYRNGK) with purity greater than 95%. The peptide was emulsified in incomplete Freund’s adjuvant (IFA; InvivoGen, France) supplemented with 400 mg/mL of Mycobacterium tuber-culosis to lead to complete Freund’s adjuvant (CFA), and injected subcutaneously near the base of the tail in a 200-μL volume. Immediately after immunization and 2 days later, mice were intraperitoneally injected with 300 ng/kg of pertussis toxin from Bordetella pertussis bac-teria (PTX; Sigma-Aldrich, Saint-Quentin Fallavier, France) in a 200-μL volume. The control animals (CFA-mice) were similarly injected with CFA free of MOG35-55 peptide and pertussis toxin.
Six mice (MOG-M1 to MOG-M6) were immunized at day 0. Recordings were performed at days 14 and 28, i.e., 2 and 4 weeks after immunization, and compared to those of six age-matched control animals (CFA-M1 to CFA-M6). All mice were daily observed for clinical signs. The references used to assess the clinical scores of EAE mice, assigned by an observer who was blinded to the treatment, were the flaccidity or paralysis of the tail (first sign), and the drag of the hip without limb paralysis (second sign) or associated with hind-limb (third sign) and then fore-limb (forth sign) paralysis.
In vivo electrophysiology
Recordings were performed by means of a minimally in-vasive electrophysiological method. The principle is to electrically stimulate a nerve trunk and to record, in re-turn, the compound action potential (CAP) resulting from the activity of all fibers composing the stimulated
nerve (“sensory nerve recordings”) or muscle (“motor nerve recordings”).
After being weighed, each mouse was placed in an anesthesia-induction chamber in which a mixture of oxygen (0.4 L/min via an oxygen extractor), air (0.2 L/min via an air condenser), and isoflurane (AErrane®, Baxter S.A., Lessines, Belgium; 2.0–2.5% via an anesthetic diffuser) was diffused. When the mouse was asleep, it was taken out of the chamber and set on a plate heated using water circulation (via a T/ PUMP). Its muzzle was positioned at the level of a mask which continuously delivered the anesthetic gas mixture to keep the animal asleep, and its two hind-limbs were fixed by adhesive tape (Fig.1). If necessary, the percentage of isoflur-ane was adjusted to maintain the isoflur-anesthesia. The animal temperature was measured using a digital thermometer equipped with a rectal probe.
The electrical stimulations were delivered to either the caudal or sciatic nerve by two stimulators (A395, World Pre-cision Instruments, Sarasota, FL, USA) via two non-polarizable Ag/AgCl external electrodes, the anode (RC4, World Precision Instruments) and the cathode (RC3, World Precision Instruments), directly affixed to the skin of the mouse. A medical gel (Polaris II) was used to improve the
contact, i.e., the electrical conduction, between the electrodes and the skin. For sensory nerve recordings, the electrodes were located at the distal portion of the mouse tail, the anode being in the most distal position and the cathode about 1 cm from the anode. For motor nerve recordings, the anode was placed at the level of the ankle of the right or left hind-limb studied, and the cathode at the base of the tail (Fig.1).
The CAP [compound nerve action potential (CNAP) and compound muscle action potential (CMAP) for sensory and motor nerve recordings, respectively], which propagated in the stimulated nerve, was collected by means of two detection electrodes E1 and E2 (MF3.OE.1F35.12, Comepa) which were very fine nee-dles inserted (i) in the proximal part of the tail, the elec-trode E2 being in the most proximal position and the electrode E1 about 1 cm from the electrode E2 (sensory nerve recordings); (ii) in the distal part of the tail, the electrode E2 being in the most distal position and the electrode E1 about 1 cm from the electrode E2 (motor nerve recordings from the tail muscle); and (iii) in the right or left hind-limb (motor nerve recordings from the plantar muscle). These electrodes were connected to an amplifier (Disa EMG 14C13, Sklovlunde) to increase the
Fig. 1 Traces of compound nerve action potential (CNAP) and compound muscle action potential (CMAP) for sensory and motor nerve recordings, respectively, evoked by nerve stimulation (upper), and corresponding locations of stimulation and detection (red arrows) electrodes (lower)
CAP amplitude, and then to a“hum bug” (Quest Scien-tific) to eliminate the sinusoidal noises that are inherent to electrophysiological recordings. Finally, a ground elec-trode was placed between the cathode and the detection electrodes for sensory and motor nerve recordings from the tail, or below the cathode for motor nerve recordings from the right or left hind-limb (Fig.1).
The multiple evaluation of sensory and motor nerve excitability properties was performed using the Qtrac© software (H. Bostock, Institute of Neurology, London, U.K.). By means of a digital-to-analog converter (DAQ2000, Iotech), this software allowed the delivery of the stimulation sequences and, in return, managed the recordings (at a sampling frequency of 10 kHz) and ana-lysis of the CNAP and CMAP collected from the stimu-lated nerve and muscle, respectively (Fig.1). It should be emphasized that, under these conditions, the CAP in re-sponse to a single stimulation was biphasic, i.e., a posi-tive phase followed by a negaposi-tive one or a negaposi-tive phase followed by a positive one, according to the rela-tive positions of the two electrodes, since it represented the potential difference between the two detection elec-trodes, i.e., E1-E2. Its amplitude was measured as the ab-solute difference between the maxima of these two phases (Fig.1).
Protocols, data analyses, and statistics
The stimulation protocol (“QTracS” program) lasted a few minutes and consisted in establishing the stimulus-response curve, i.e., the relationship between the CAP amplitude and the stimulation intensity, as exemplified for the CNAP in Fig. 2. Firstly, the CAP amplitude was measured as a function of the intensity of the stimula-tion (electric current) of 1-ms durastimula-tion which, starting from 0, was gradually increased by steps of 3% of its
maximum value (i.e., 2 mA, each of the two stimulators delivering a maximum of 1 mA), and manually until a maximum CAP amplitude was obtained. Secondly, a stimulus-response curve was generated automatically by the program, from which four parameters characteristic of sensory nerve or neuromuscular excitability were esti-mated (“QTracP” program). These parameters included (1) the maximal CAP amplitude (CAPmax), which depended on the number of muscle and nerve fibers responding to stimulation; (2) the stimulation intensity necessary to generate a CAP with an amplitude equal to 50% of its maximum value (SI-50%), which depended on the excitability threshold of fibers; (3) the slope of the stimulus-response curve (Slope), which depended essen-tially on the passive membrane properties of fibers; and (4) the time between the stimulus onset and the first peak amplitude of CAP (Latency), which depended on the CAP propagation/transmission velocity. Therefore, changes in these parameters gave information mainly on the density and functional state (activation) of voltage-gated sodium (NaV) and potassium (KV) channels, as well as on the passive membrane properties of nerve fi-bers linked, in particular, to the presence or absence of a myelin sheath surrounding the axons.
Data are presented as means ± standard deviations (S.D.) of at least 5 (n) different mice (6 and 5–6 animals 2 and 4 weeks after immunization, respectively). Differ-ences between values were tested using the parametric two-tailed Student’s t test (either paired samples for comparison within a single population or unpaired sam-ples for comparison between two independent popula-tions), and the one- or two-way analysis of variance (ANOVA for comparison between the means of inde-pendent populations) or the non-parametric Mann-Whitney U test, depending on the equality of variances
Fig. 2 Stimulus-response curves (i.e., CNAP in response to nerve stimulations of increasing intensity) and derived excitability parameters (left). Relationships between the CNAP amplitude and the stimulation intensity delivered manually (in pink) and then automatically (in green) by the program (right)
estimated using the Lilliefors’ test. Differences between results are considered to be statistically significant for a P value of less than or equal to 0.05.
Evaluation of myelin thickness through transmission electronic microscopy
Under anesthesia with ketamine and xylazine (75 mg/kg and 10 mg/kg, respectively, intraperitoneal), mice were perfused with modified Karnovsky fixative solution con-taining 2.5% glutaraldehyde and 2% paraformaldehyde in 0.1 M sodium phosphate buffer solution (pH 7.4). The muscles of the right hind leg were dissected and the right sciatic nerve was collected. Samples were post-fixed in a solution of 1% osmium tethoxide, at 4 °C, followed by immersion in a 5% aqueous uranyl acetate solution at room temperature. After dehydration in alco-hol, samples were immersed in propylene oxide and then included in Spur resin. Samples were sectioned (semi-thin sections—15 μM) in an ultra-microtome (Reichert Ultra Cut®) and stained with 1% toluidine blue solution. Subsequently, ultra-thin sections were cut (60 nm), col-lected them on 200 mesh copper grid (Sigma®), and the contrast was obtained using 4% uranyl acetate solution
and 0.4% aqueous lead citrate solution [40]. The grids were examined in the Jeol 1010 transmission electron microscope (Department of Anatomy at the University of São Paulo) and quantification was performed using ImageJ software (NIH/EUA).
Results
Clinical scores of EAE and control mice
The clinical scores of MOG- and CFA-mice were deter-mined every day after immunization, for 6 weeks, to de-scribe the motor symptoms that occurred in the progression of EAE (Fig.3).
As expected, the mean clinical score increased from 0.0 to more than 2.5 within about 18 days after immunization for MOG-mice while it remained null for all CFA-mice. It is worth noting that a quite large clin-ical score variability occurred between EAE mice.
Mouse body weight and temperature
Significant decreased body weight were observed in MOG-mice compared to CFA-animals, 2 weeks (P = 0.001) and, although less pronounced, 4 weeks (P = 0.027) after immunization (Fig. 4). However, compared
Fig. 3 Individual clinical scores of 6 EAE mice (MOG-M1 to MOG-6) and control mice (CFA-M1 to CFA-6) as a function of time after immunization at day 0. The red arrows indicate the days of electrophysiological recordings (i.e., 2 and 4 weeks) after immunization
to initial body weight measurements performed the day of immunization (day 0), i.e., 19.58 ± 0.43 (n = 6), EAE mice lost weight 2 weeks after immunization (P = 0.0002) and then gained weight 4 weeks after immunization (P = 0.435). On the contrary, control animals homogeneously gained weight (P < 0.013), compared to initial body weight measurements performed at day 0, i.e., 19.63 ± 0.28 (n = 6).
No significant change (P ≥ 0.492) in body temperature was detected between EAE and control mice, whatever the number of weeks after immunization (Fig. 4).
Sensory nerve excitability properties (CNAP recordings)
As shown in Fig.5, a significant decrease of the maximal CNAP amplitude (P ≤ 0.012) and of the slope of the stimulus-response curve (P ≤ 0.008), as well as a signifi-cant increase in the time between the stimulus onset and the first peak amplitude of CNAP (P ≤ 0.035), were observed in MOG-mice, compared to CFA-animals, at any given time-point. This indicates a lower propagation velocity of CNAP in EAE than in control animals.
These changes in the four parameters derived from the stimulus-response curve are exemplified in Fig.6 by CNAP recordings obtained from individual MOG- and CFA-mice, 4 weeks after immunization.
Similarly, low propagation velocity of CNAP (increased Latency) and sensory nerve hyperexcitability (decreased SI-50%) in EAE mice, compared to control animals, are exemplified in Fig.7by CNAP recordings obtained from individual MOG- and CFA-mice, 2 weeks after immunization.
Motor nerve excitability properties (CMAP recordings)
The CMAP recorded at the tail and plantar muscles of EAE mice was generally followed by a second CMAP (of
reduced amplitude), as illustrated in Fig.8, 4 weeks after immunization.
The second CMAP was detected in 50% EAE mice (3/6 animals) 2 weeks after immunization while, 4 weeks after immunization, it was systematically ob-served in 100% EAE mice (5/5 animals). This second CMAP never occurred in control mice (6/6 and 6/6 animals, 2 and 4 weeks after immunization, respect-ively). As already stated in the “Materials and methods” section, the CMAP in response to a single
stimulation consisted of a positive phase followed by a negative one (as exemplified by tail muscle record-ings) or a negative phase followed by a positive one (as exemplified by plantar muscle recordings), accord-ing to the relative positions of the two detection electrodes.
Expressing the CMAP amplitude as a percentage of the first CMAP amplitude revealed a high inter-individual variability in the second CMAP amplitude (Fig.9a–b), as already observed for the EAE mouse clin-ical scores (see Fig. 3). Indeed, establishing the relation-ship between the second CMAP amplitude and the EAE mouse clinical scores revealed a good correlation (r2 ≥ 0.677) between these two parameters (Fig.9c). It is thus likely that the individual variability in both the presence and amplitude of the second CMAP reflected that of clinical scores.
In addition to the presence of a second CMAP in EAE mice, a significant decrease of the stimulation intensity necessary to generate a CMAP with an amplitude equal to 50% of its maximum value (P ≤ 0.038) was detected in MOG-mice, compared to CFA-animals, at any given time-point (Table 1, Supplementary Figures 1and 2), as observed for the CNAP (see Fig. 5). However, the delay between the two CMAPs, measured as the time between their first peak amplitude, was constant, i.e., between 3.5
Fig. 4 Body weight (left) and temperature (right) of EAE (MOG) mice, compared to control (CFA) animals, 2 (n = 6) and 4 (n = 5–6) after immunization. *P = 0.027 and **P = 0.001
Fig. 5 Maximal CNAP amplitude (CNAPmax), stimulation intensity necessary to generate a CNAP with an amplitude equal to 50% of its maximum value (SI-50%), slope of the stimulus-response curve (Slope), and time between the stimulus onset and the first peak amplitude of CNAP (Latency) in EAE (MOG) mice, compared to control (CFA) animals, 2 (n = 3–5) and 4 (n = 5–6) weeks after immunization. *P ≤ 0.035, **P ≤ 0.008, and ***P ≤ 0.0005
Fig. 6 Example of increased Latency and decreased CNAPmax, Slope, and SI-50% in an EAE (MOG) mouse, compared to a control (CFA) animal, 4 weeks after immunization
and 4 ms (see Fig.8), and no significant difference of the time between the stimulus onset and the first peak amp-litude of the first CMAP (P ≥ 0.292) was detected be-tween EAE and control mice (Table 1, Supplementary Figures1and2).
Additionally, and as already observed for the CNAP (see Fig.5), a significant decrease of the maximal CMAP amplitude (P ≤ 0.039) and of the slope of the stimulus-response curve (P ≤ 0.048) was observed in MOG-mice,
compared to CFA-animals, at any given time-point (Table1, Supplementary Figures1and2).
Myelin sheath thickness of EAE and control mice
To verify the alterations in the myelin contend in-duced in the EAE model, the morphology of the dis-tal portion of the sciatic nerve was analyzed by transmission electron microscopy on the 2nd (peak of the disease) and 4th weeks after immunization. The
Fig. 7 Low propagation velocity of CNAP (increased Latency) and sensory nerve hyperexcitability (decreased SI-50%) in EAE (MOG) mice, compared to control (CFA) animals, are exemplified by CNAP recordings showing (1)“control-like” (i.e., unaffected), (2) “slower”, (3) “very slower”, and (4) “very very slower” conducting nerve fibers in a MOG-mouse, compared to a CFA-animal, 2 weeks after immunization
thickness of the myelin sheath was quantified by cal-culating the g-ratio (i.e., the axon diameter divided by the fiber diameter). The results demonstrated intact fibers, with similar distribution of myelinic fibers of small and large diameters, non-myelinic fibers, and Schwann cell nuclei in the control group. No
difference in myelin sheath thickness was observed between control and EAE mice, at the peak of the disease. In contrast, a decreased sciatic nerve myelin thickness, represented by an increase in the g-ratio, was observed in the EAE group, compared to control animals, in the 4th week after immunization (Fig. 10).
Fig. 9 Expression of CMAP amplitude as a percentage of the first CMAP amplitude in individual EAE mice (MOG-M1 to MOG-6), 2 and 4 weeks after immunization (a and b), and relationship between the second CMAP amplitude and the EAE mouse clinical score (c)
Table 1 Parameters (means ± S.D.) of the stimulus-response curve determined from tail and plantar muscles in n (numbers in parentheses) EAE (MOG) mice, compared to control (CFA) animals, 2 and 4 weeks after immunization
2 weeks 4 weeks
CFA-mice (6) MOG-mice (6) CFA-mice (6) MOG-mice (5) Tail muscle CMAPmax (mV) 6.922 ± 0.773 3.893 ± 0.544* 5.992 ± 0.469 3.787 ± 0.511* SI-50% (mA) 0.222 ± 0.014 0.179 ± 0.006* 0.224 ± 0.015 0.166 ± 0.013* Slope 5.226 ± 0.451 3.258 ± 0.515* 4.977 ± 0.238 3.982 ± 0.112* Latency (ms) 3.854 ± 0.067 3.703 ± 0.104 3.987 ± 0.130 3.818 ± 0.075 Plantar muscle CMAPmax (mV) 6.075 ± 0.650 3.204 ± 0.513** 5.635 ± 0.588 3.011 ± 0.522* SI-50% (mA) 0.278 ± 0.023 0.174 ± 0.010** 0.320 ± 0.014 0.231 ± 0.013** Slope 6.261 ± 0.650 3.559 ± 0.405* 5.756 ± 0.337 4.051 ± 0.206** Latency (ms) 3.044 ± 0.201 3.625 ± 0.223 3.286 ± 0.147 3.346 ± 0.205
CMAPmax maximal CMAP amplitude, SI-50% stimulation intensity necessary to generate a CMAP with an amplitude equal to 50% of its maximum value, Slope slope of the stimulus-response curve,Latency time between the stimulus onset and the first peak amplitude of CMAP. *P ≤ 0.048 and **P ≤ 0.007
Discussion
The results of this study can be summarized as follow: (i) a large inter-individual clinical score variability between EAE mice, (ii) a difference in body weight between EAE and control mice, (iii) modifications of the peripheral sen-sory nerve excitability parameters, and (iv) modifications of the neuromuscular excitability parameters.
The large inter-individual variability in clinical score between EAE mice may be attributed to differences in the susceptibility to EAE, as previously observed even in genetically identical animals [41]. Despite this variability in the intensity of the clinical score among the animals, all immunized mice developed some degrees of motor impairment characteristic of the disease, while none of the control animals showed this symptom.
We observed that EAE mice first lost and then pro-gressively gained weight while control animals homoge-neously gained weight, compared to initial body weight measurements performed at day 0. As a consequence, a difference in body weight between EAE and control mice, which is less marked as the number of weeks in-creased after immunization, was detected. These obser-vations may be related to the fact that EAE mice had moving difficulties because of hind-limb paralysis and, therefore, did less exercise than control mice and that EAE animals showing a more than 2.0 clinical score were fed with highly nutritious food.
The results obtained from CNAP and CMAP record-ings strongly suggest that the peripheral sensory nerve and neuromuscular excitability properties of EAE mice are markedly modified compared to those of control ani-mals, 2 and 4 weeks after immunization. These modifi-cations are not due to differences in body temperature since no significant change in this parameter was de-tected between EAE and control mice, whatever the number of weeks after immunization. They consist in a significant decrease of maximal CNAP and CMAP am-plitudes, of the stimulation intensity necessary to
generate a CNAP or CMAP with an amplitude equal to 50% of its maximum value, and of the slope of stimulus-response curves at any given time-point. In addition, a significant increase in the time between the stimulus on-set and the first peak amplitude of CNAP was observed in MOG-mice compared to CFA-animals, at any given time-point, indicating a lower propagation velocity of the electrical transmission of the action potential in EAE than in control animals. In contrast, no significant differ-ence of the time between the stimulus onset and the first peak amplitude of the first CMAP was detected between EAE and control mice. This strongly suggests that the chemical transmission of the action potential at the neuromuscular junction is not delayed. These electro-physiological abnormalities are similar to those previ-ously reported in the ulnar and sural sensory nerves of some MS patients diagnosed according to the criteria of Poser Scale [28] and in superficial radial sensory axons of a few patients with relapsing-remitting MS [29]. They are also in accordance with the high fre-quency of electrophysiological abnormalities previ-ously reported in the tibial motor nerve of a selected group of MS patients [28].
The significant decrease of the stimulation intensity necessary to generate a CNAP or a CMAP with an amp-litude equal to 50% of its maximum value detected in MOG-mice, compared to CFA-animals, indicates sen-sory and motor nerve hyperexcitability in EAE mice compared to control animals. In agreement, membrane hyperexcitability of sensory neurons of dorsal root gan-glia isolated from MOG35-55-induced EAE mice was pre-viously reported [23]. Besides, the presence of a second CMAP in EAE mice may also express membrane hyper-excitability. However, the fact that the delay between the two CMAPs, measured as the time between their first peak amplitude, was constant strongly suggests that this second CMAP may also result from the activity of slow conducting motor nerve fibers, as expected if a
Fig. 10 Peripheral nerve myelin sheath of EAE (MOG) mice, compared to control (CFA) animals, 2 and 4 weeks after immunization. Data represent the mean ± S.D. of 48 randomly chosen fibers per group, each constituted of 3 mice (i.e., 16 fibers per mouse). *P < 0.0001
demyelinating process occurs in the peripheral nervous system of EAE mice. Under these conditions, the first CMAP would be due to unaffected axons, whereas the second one would correspond to partially demyelinated axons. This is in agreement with the absence of detec-tion of time difference between the stimulus onset and the first peak amplitude of the first CMAP in EAE mice, compared to control animals. Moreover, the recordings of multiple (at least 4) CNAP peaks may reflect not only the decrease of both the propagation velocity and the SI-50% parameter of CNAP but also various populations of sensory nerve fibers having different CNAP conduction speeds in EAE mice. This is expected if a demyelinating process occurred in the peripheral nervous system of EAE mice.
To support the occurrence of a peripheral demyelinat-ing process, we investigated the alterations of sciatic nerve myelin contend by transmission electron micros-copy, a methodology previously used as a parameter of peripheral myelin analysis for evaluation of neuroprotec-tive therapies evaluation [42]. Although the widely ex-pression of myelin peptides as MBP and MOG in the CNS is remarkable [43–46], these proteins are also expressed in the PNS [47–50], the thymus, and the spleen [51, 52]. These proteins can therefore be a target of autoantibodies attack, contributing to demyelination at peripheral level and, as a consequence, to peripheral sensory and motor alterations.
Conclusion
In conclusion, the main modifications of the peripheral sensory nerve and neuromuscular excitability are (i) a membrane hyperexcitability likely related to membrane depolarization and (ii) the presence of slow conducting sensory and motor nerve fibers due to a demyelinating process occurring in the PNS of EAE mice. These modi-fications are of great interest since the loss of immuno-logical self-tolerance to MOG in EAE animal model or in patients with MS is generally attributed to the re-stricted expression of this antigen in the immunologic-ally privileged environment of the CNS.
Supplementary information
Supplementary information accompanies this paper athttps://doi.org/10. 1186/s12974-020-01936-9.
Additional file 1: Supplementary Figure 1. Parameters of stimulus-response curves determined from tail muscle recordings in EAE (MOG) mice compared to control (CFA) animals. Supplementary Figure 2. Pa-rameters of stimulus-response curves determined from plantar muscle re-cordings in EAE (MOG) mice compared to control (CFA) animals.
Abbreviations
ANOVA:Analysis of variance; ARRIVE: Animal Research: Reporting of In Vivo Experiments; CAP: Compound action potential; CAPmax: Maximal CAP amplitude; CFA: Complete Freund’s adjuvant; CMAP: Compound muscle
action potential; CNAP: Compound nerve action potential; CNS: Central nervous system; EAE: Experimental autoimmune encephalomyelitis; IFA: Incomplete Freund’s adjuvant; KVchannel: Voltage-gated potassium
channels; Latency: Time between the stimulus onset and the first peak amplitude of CAP; MBP: Myelin basic protein; MOG: Myelin oligodendrocyte glycoprotein; MS: Multiple sclerosis; NaVchannel: Voltage-gated sodium
channel; PNS: Peripheral nervous system; SI-50%: Stimulation intensity necessary to generate a CAP with an amplitude equal to 50% of its maximum value; Slope: Slope of the stimulus-response curve; Th1: T-helper type 1 cell; Th17: T-helper type 17 cell
Acknowledgements Not applicable.
Authors’ contributions
GP, FB, GP, BK, DS, and EB conceived and designed the experiments; NBT, ACG, and EB performed and analysed the experiments; GP and EB wrote and prepared the original draft; GP, ACG, and EB wrote and prepared the revised draft; GP, FB, GP, BK, DS, and EB reviewed and edited the paper. GP, FB, BK, and DS acquired the funding. All authors read and approved the final version of the manuscript to be published.
Funding
This research was financed by funds from the French Alternative Energies and Atomic Energy Commission (CEA, Gif sur Yvette, France), from the São Paulo Research Foundation (FAPESP, Brazil, grant number 2011/17974-2), from FAPESP—Center of Toxins, Immune Response and Cell Signaling (CeTICS, grant number 2013/07467-1), and from the Coordenação de Aperfeiçoamento de Pessoal de Nível Superior—Brasil (CAPES, Finance Code 001). Research collaboration was also supported by the IASP Developing Countries Collaborative Research Grant.
Availability of data and materials
All datasets [GENERATED/ANALYZED] for this study are included in the manuscript and the supplementary files.
Ethics approval and consent to participate
Animal experiments, reported in line with the ARRIVE (Animal Research: Reporting of In Vivo Experiments) guidelines developed in consultation with the scientific community as part of an NC3Rs initiative to improve standards of reporting the results of animal experiments, maximizing information published and minimizing unnecessary studies, were carried out on mice purchased from Janvier Elevage (Le Genest-Saint-Isle, France). The animals were hosted at the CEA animal facility and were treated in strict adherence with the European Community guidelines for laboratory animal handling and to the guidelines established by the French Council on animal care “Guide for the Care and Use of Laboratory Animals” (EEC86/609 Council Directive—Decree 2001-131). All animal experimental procedures were ap-proved by the Animal Ethics Committee of the CEA, by the French General Directorate for Research and Innovation (project
APAFIS#5973-2016070515456532v6 authorized to FB and project APAFIS#2671-2015110915123958v4 authorized to EB) and by the Butantan Institute (CEUAIB protocol number 7334170718 authorized to GP).
Consent for publication Not applicable.
Competing interests
The authors declare that they have no competing interests. Author details
1Université Paris-Saclay, CEA, Département Médicaments et Technologies
pour la Santé (DMTS), Service d’Ingénierie Moléculaire pour la Santé (SIMoS), ERL CNRS 9004, Gif-sur-Yvette, France.2Laboratory of Pain and Signaling, Butantan Institute, São Paulo, Brazil.3Université Paris-Saclay, CEA, NeuroSpin,
Gif-sur-Yvette, France.4Université Paris-Saclay, CEA, Inserm, BioMaps, Orsay,
Received: 5 May 2020 Accepted: 20 August 2020
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